For more than a century, the primary clinical identification method for bacterial infections has been plate cultures where bacteria are isolated and purified overnight using nutrient-based agar medium. Although rapid, automated instrumentation has been widely regarded as the next step towards advancing bacterial identification, these instruments still require a pure bacterial colony obtained from a plate culture and thus a lead time of at least 18-24 hours before any identification can be made. (Cherkaoui, A., et al., Comparison of two matrix-assisted laser desorption ionization-time of flight mass spectrometry methods with conventional phenotypic identification for routine identification of bacteria to the species level. J Clin Microbiol, 2010. 48(4): p. 1169-75. Holland, R. D., et al., Rapid identification of intact whole bacteria based on spectral patterns using matrix-assisted laser desorption/ionization with time-of-flight mass spectrometry. Rapid Communications in Mass Spectrometry, 1996. 10(10): p. 1227-1232.) Likewise, molecular diagnostics such as polymerase chain reaction require pure bacterial colonies and hours of processing. (Meng, J., et al., Polymerase chain reaction for detecting Escherichia coli O157: H7. Int J Food Microbiol, 1996. 32(1-2): p. 103-13.) As a result, rapid screens for clinically-relevant bacterial species have become an attractive alternative option for hospitals looking for more rapid point-of-care diagnostics. A rapid screen for Pseudomonas aeruginosa and other clinically-relevant bacteria would allow doctors to promptly switch from broad-spectrum antibiotics to specific directed therapies, lowering hospital expenditures, minimizing drug resistance, and improving patient care outcomes. (Trenholme, G. M., et al., Clinical impact of rapid identification and susceptibility testing of bacterial blood culture isolates. J Clin Microbiol, 1989. 27(6): p. 1342-5.)
Additionally, the ability to monitor the effect antibiotics have on bacteria is important for infection control. The conventional approach to determining antibiotic efficacy requires the creation of culture plates with the antibiotic cocktail of choice at a series of concentrations. (J. M. Andrews, J. Antimicrob Chemother, 2001, 48, 5-16.) After culturing (for 24 hours or longer, depending on the strain), the plates are visually inspected for growth. At a certain concentration, known as the minimum inhibitory concentration (MIC), no bacterial growth is observed. This concentration is then used to design an antibiotic schedule for the patient. This effective approach suffers from the use of large amounts of reagents required to produce the culture plates. Furthermore, these screens only measure the effectiveness of the antibiotic against planktonic cell growth; not removal of biofilms, which are commonly associated with infections and significantly more difficult to treat. (P. K. Singh, A. L. Schaefer, M. R. Parsek, T. O. Moninger, M. J. Welsh and E. P. Greenberg, Nature, 2000, 407, 762-764. C. F. Schierle, M. De la Garza, T. A. Mustoe and R. D. Galiano, Wound Repair Regen, 2009, 17, 354-359.)
An alternative is coupling microfluidics, to grow the bacteria, with antibiotic screens. Kim et al. (2012) utilized a microfluidic system to simultaneously expose biofilms of Escherichia coli to eight different concentrations of antibiotics on a single chip. (J. Kim, M. Hegde, S. H. Kim, T. K. Wood and A. Jayaraman, Lab Chip, 2012, 12, 1157-1163.) The smaller volumes, inherent in microfluidic devices, along with the ability to produce multiple concentration gradients provided a faster, cheaper alternative to current antibiotic susceptibility tests. By flowing antibiotics over the grown biofilms, researchers more closely simulated in vivo conditions. Many current microfluidic studies determine biofilm viability based on the presence of fluorescent proteins during exposure to antibiotics. (J. Kim, H. D. Park and S. Chung, Microfluidic approaches to bacterial biofilm formation, Molecules, 2012, 17, 9818-9834. K. P. Kim, Y. G. Kim, C. H. Choi, H. E. Kim, S. H. Lee, W. S. Chang and C. S. Lee, Lab Chip, 2010, 10, 3296-3299.) While these methods are certainly robust and promising, the fluorescent signal requires expensive optical equipment and genetically modified bacteria or selective labels. (L. Richter, C. Stepper, A. Mak, A. Reinthaler, R. Heer, M. Kast, H. Bruckl and P. Ertl, Lab Chip, 2007, 7, 1723-1731. H.-Y. N. Holman, R. Miles, Z. Hao, E. Wozei, L. M. Anderson and H. Yang, Anal Chem, 2009, 81, 8564-8570. Y. Yawata, K. Toda, E. Setoyama, J. Fukuda, H. Suzuki, H. Uchiyama and N. Nomura, J Biosci Bioeng, 2010, 110, 130-133. Y. Yawata, K. Toda, E. Setoyama, J. Fukuda, H. Suzuki, H. Uchiyama and N. Nomura, J Biosci Bioeng, 2010, 110, 377-380.) A cheaper and easier method of determining the relative amount of live cells in a biofilm under exposure to antibiotics can be achieved by monitoring the electrochemical response of the system. Robust bacterial biofilms produce a plethora of molecules that promote communication, defend the colony, and cause infection. (M. B. Miller and B. L. Bassler, Annu Rev Microbiol, 2001, 55, 165-199. M. D. P. Willcox, H. Zhu, T. C. R. Conibear, E. B. H. Hume, M. Givskov, S. Kjelleberg and S. A. Rice, Microbiology, 2008, 154, 2184-2194. G. W. Lau, D. J. Hassett, H. Ran and F. Kong, Trends Mol Med, 2004, 10, 599-606.) Of interest are molecules that provide information about the condition of the biofilm, which can be detected by electrochemical methods.